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Detection of Thiol Modifications by Hydrogen Sulfide E. Williams*, S. Pead*, M. Whiteman†, M.E. Wood{, I.D. Wilson*, M.R. Ladomery*, T. Teklic}, M. Lisjak}, J.T. Hancock*,1 *Faculty of Health and Applied Sciences, University of the West of England, Bristol, United Kingdom † University of Exeter Medical School, University of Exeter, Exeter, United Kingdom { Biosciences, University of Exeter, Exeter, United Kingdom } Faculty of Agriculture, University of Osijek, Osijek, Croatia 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Hydrogen Sulfide Acts as a Signal in Cells 3. Modification of Thiols by Signaling Molecules 4. Identification of Modified Thiols by Other Methods 5. Experimental Protocols 6. Caenorhabditis elegans as a Model Organism 7. Growth of C. elegans 8. Treatment of Samples with H2S 9. Estimation of Toxicity of H2S Compounds 10. Treatment of Samples with Thiol Tag 11. Isolation and Analysis of Modified Proteins 12. Estimation of Protein Concentrations in Samples 13. Further Analysis and Identification of Modified Proteins 14. Concluding Remarks References

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Abstract Hydrogen sulfide (H2S) is an important gasotransmitter in both animals and plants. Many physiological events, including responses to stress, have been suggested to involve H2S, at least in part. On the other hand, numerous responses have been reported following treatment with H2S, including changes in the levels of antioxidants and the activities of transcription factors. Therefore, it is important to understand and unravel the events that are taking place downstream of H2S in signaling pathways. H2S is known to interact with other reactive signaling molecules such as reactive oxygen species (ROS) and nitric oxide (NO). One of the mechanisms by which ROS and NO have effects in a cell is the modification of thiol groups on proteins, by oxidation or S-nitrosylation, respectively. Recently, it has been reported that H2S can also modify thiols. Here we report a method for the determination of thiol modifications on proteins following Methods in Enzymology ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2014.11.026

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2015 Elsevier Inc. All rights reserved.

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the treatment with biological samples with H2S donors. Here, the nematode Caenorhabditis elegans is used as a model system but this method can be used for samples from other animals or plants.

1. INTRODUCTION Hydrogen sulfide (H2S) is now known to be an important metabolite involved in the control of cellular function in a range of organisms, including both animals and plants. It has been suggested that it is considered as part of the suite of gasotransmitters in organisms (Wang, 2002, 2003), despite the fact that is very toxic and a potent inhibitor of mitochondrial function through inhibition of Complex IV (Dorman et al., 2002). In mammals, H2S has been shown to be involved in the control of vascular tone (Liu et al., 2012) and to have a role in related diseases, while in plants H2S has been shown to be involved in alleviation of numerous stress responses as well as being involved in the control of normal physiological functions (Calderwood and Kopriva, 2014), such as the regulation of stomatal apertures (Garcı´a-Mata & Lamattina, 2010; Lisjak et al., 2010), flower senescence (Zhang et al., 2011), and root development (Lin, Li, Cui, Lu, & Shen, 2012). Therefore, the downstream events following the exposure of cells to H2S, either from the generation of H2S by cells themselves or from the environment, are important to determine. Cells can be exposed to H2S from a variety of sources. From the environment, cells from both animals and plants can come into contact with H2S which has been released from volcanoes, for example, while evolution may have been driven by the H2S from thermal vents (Martin, Baross, Kelley, & Russell, 2008). Bacteria can make H2S (Clarke, 1953), so exposing higher organisms to this gas. Even during warfare, humans have been exposed to this gas (Szinicz, 2005), highlighting its toxic nature. Endogenously, in plants H2S can be generated by desulfhydrases (Alvarez, Calo, Romero, Garcia, & Gotor, 2010), while in animals the gas may be made by 3-mercaptopyruvate sulfurtransferase (3-MST), cystathionine-γ-lyase, or cystathionine-β-synthase (Prabhakar, 2012). It is important to note here that H2S can be removed by cells too. In plants, the enzyme O-acetylserine(thiol) lyase (Tai & Cook, 2000; Youssefian, Nakamura, & Sano, 1993) can reduce H2S levels, and in animals it has been shown that H2S can be used by mitochondria as a source of reducing equivalents, leading to the generation of

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ATP (Bouillaud, Ransy, & Andriamihaja, 2013), and the removal of H2S from the cells. Therefore, with enzymes able to generate and remove H2S, its use as a signal does not seem a surprise.

2. HYDROGEN SULFIDE ACTS AS A SIGNAL IN CELLS In humans, H2S is known to be involved in a range of physiological events, often centered around the regulation of the cardiovascular system (Liu et al., 2012). Furthermore, work in humans has shown that treatment with H2S has been shown to alleviate the onset and symptoms of several diseases (Ahmed, 2013; Al-Magableh, Ng, Kemp-Harper, Miller, & Hart, 2013; Nagpure & Bian, 2013; Wang, 2013), among which are diabetes (Whiteman et al., 2010), atherosclerosis (Mani et al., 2013; Xu, Liu, & Liu, 2014), and vascular inflammation (Liu et al., 2013). In plants, H2S has been shown to alleviate the stress responses to a range of challenges, including to heavy metals (Ali et al., 2014; Li, Wang, & Shen, 2012; Zhang et al., 2008, 2010), temperature (Li, Ding, & Du, 2013; Li, Gong, Xie, Yang, & Li, 2012; Stuiver, De Kok, & Kuiper, 1992), and osmotic (Zhang et al., 2009) and oxidative stress (Shan et al., 2011; Zhang et al., 2008). The interaction of H2S with oxidative stress is particularly pertinent here, as both are involved in a range of stress responses. H2S has been shown to have interactions with both the metabolism of reactive oxygen species (ROS) and nitric oxide (NO), as discussed previously (Hancock & Whiteman, 2014). Treatment with H2S has an influence on the enzymes which produce ROS and NO, as well as the mechanisms to remove these compounds, that is, on antioxidants. For example, in plants, treatment with H2S can cause alterations in antioxidant levels following salt stress (Lisjak, Teklic, Wilson, Whiteman, & Hancock, 2013), and postharvest storage can be enhanced, again mediated by the alteration of levels of antioxidants (Hu et al., 2012). Glutathione levels too have been reported to be increased in plant cells following treatment with H2S (De Kok, Bosma, Maas, & Kuiper, 1985), so increasing the capacity of a cell to tolerate oxidative stress conditions. Therefore, it can be seen that organisms from bacteria to humans are exposed to H2S and indeed make it for positive reasons. H2S can be used as a signaling molecule, although at high concentrations it is toxic. It can be used for mitochondria ATP generation (Bouillaud et al., 2013), and it is known to have effects on other signaling pathways, especially those involving ROS and NO (Hancock & Whiteman, 2014). Determining the biomolecules that H2S interacts with is therefore important.

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3. MODIFICATION OF THIOLS BY SIGNALING MOLECULES Therefore, as can be seen from above, H2S can impinge on the signaling which involves either ROS or NO, or indeed both (Hancock & Whiteman, 2014). One of the convergence points of this type of signaling is the modification of thiol groups in proteins (Chouchani, James, Fearnley, Lilley, & Murphy, 2011). It has been known for some time that thiol groups can undergo a range of chemistries, as indicated in Fig. 1. The –SH group may interact to form disulfides, an important element in the folding and three-dimensional structure of proteins. Alternatively, the –SH group may be sequentially oxidized to the sulfenic acid group, the sulfinic acid group to the sulfonic acid group. The formation of the sulfenic acid, which is relatively unstable, has been shown to be reversible, or to partake in other reactions (Biteau, Labarre, & Toledano, 2003) and so this could be a good way to control the activity of a protein, akin to phosphorylation and dephosphorylation. Furthermore, the next higher oxidation state of the thiol, that is, the sulfinic acid is also thought to be able to be reversed (Biteau et al., 2003), again suggesting that its involvement in signaling is suitable. The highest oxidation state, the sulfonic acid, is thought to be irreversible. However, in tyrosine phosphatases a further chemistry of the thiol has been suggested, that is, a ring structure has been proposed, allowing the reversion of the thiol back to its original state (Salmeen et al., 2003). In the presence of NO, the thiol will partake in a reaction which leads to what is referred to as S-nitrosylation, which again has been shown to be Disulfide bond formation (may stabilize proteins)

–SH (protein thiol)

Oxidation (higher oxidation forms as hydrogen peroxide concentration increases) S-nitrosylation (with nitric oxide) Glutathionation (possible with glutathione)

Sulfhydration (in the presence of hydrogen sulfide)

Figure 1 Some of the thiol modifications which can take place in proteins in the presence of a range of chemicals, including H2S.

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reversible and so a good way to control the function of a protein (Hess, Matsumoto, Kim, Marshall, & Stamler, 2005). To assess the extent of S-nitrosylation of proteins, assays such as that dubbed the “biotin switch” assay have been developed ( Jaffrey, Erdjument-Bromage, Ferris, Tempst, & Snyder, 2001) and extensively used (Forrester, Foster, Benhar, & Stamler, 2009; Haldar & Stamler, 2013; Nakamura et al., 2013; Zhang, Keszler, Broniowska, & Hogg, 2005). This assay is further discussed below. A different approach was used to assess the proteins which may be oxidized by hydrogen peroxide (H2O2) in plants (Hancock et al., 2005), and it is this rationale which is proposed here (see Fig. 2). A similar approach has also been used with animal cells (Baty, Hampton, & Winterbourn, 2005). In this assay, free thiols are reacted with a labeled iodoacetamide, in this case 50 iodoacetamide fluorescein (IAF). Once separated on an acrylamide gel by electrophoresis, the reacted proteins can be visualized by the fluorescent nature of the tag. A separate, but identical sample, is pretreated with H2O2 and then subsequently reacted with IAF, separated by acrylamide electrophoresis and visualized. Any protein spots deemed to have disappeared when compared to the original sample are assumed to have reacted with the H2O2 before the treatment with IAF; hence, they appear to be missing in the electrophoretic analysis. When this approach was used

Cellular effects S

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Identification using 2-D electrophoresis and mass spectrometry

SO3H

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Figure 2 A schematic explaining the rationale behind the approach used to identify thiol groups in proteins which may react with H2O2 or H2S.

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previously, several proteins were identified as potentially reacting with H2O2 in cells, including glyceraldehyde 3-phosphate dehydrogenase (GAPDH), alcohol dehydrogenase, and SAM synthase. With a subsequent focus on GAPDH, it could be shown in vitro that this enzyme did indeed react in a reversible manner with H2O2 and interestingly also NO (Hancock et al., 2005), showing the validity of this approach. Further work on the reaction of GAPDH with H2O2 or NO showed that on modification the enzyme moved to the nucleus where it appeared to have a function in controlling gene expression. Hence, the identification of GAPDH as a target for H2O2 facilitated the elucidation of a whole new role for this enzyme, which was previously thought to be well characterized (Tristan, Shahani, Sedlak, & Sawa, 2011). Clearly, therefore, the identification of thiol modifications of proteins, by ROS, NO, or H2S, can lead to a further understanding of both the regulation and roles of proteins in cells.

4. IDENTIFICATION OF MODIFIED THIOLS BY OTHER METHODS One of the most well-used methods for identification of thiol modifications of proteins is the so-called biotin switch assay as mentioned above (Forrester et al., 2009; Zhang et al., 2005). This assay was first suggested over 10 years ago ( Jaffrey et al., 2001) and has been used extensively since that time to determine the modification of thiol groups by NO. The rationale behind the assay is thiol groups may have reacted with NO, perhaps in a physiological setting, but not all thiols would have reacted. So remaining free thiols are reacted with methyl-methane thiosulfonate (MMTS), which will readily react with protein thiols. This leaves all the thiols either S-nitrosylated, because they had previously reacted with NO, or S-methylthionated, as they had reacted with the MMTS. Any thiols which had been S-nitrosylated are then reduced with ascorbate, removing the NO group and reforming the –SH group. This reaction is assumed to be specific as thiols which had previously been oxidized to sulfenic acid or sulfinic acid are not rereduced, and any which have previously been glutathionated are also unaffected. So the previously nitrosylated thiols are then reacted with biotin-HPDP, so labeling the thiols for identification, using avidinhorseradish peroxidase. The use of this assay, the pitfalls, and suggested modifications and controls have been previously discussed (Forrester et al., 2009), but it is an assay which has caused some controversy and needs to be undertaken with care (Kovacs & Lindermayr, 2013).

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In the biotin switch assay, there is a requirement for the –SNO (S-nitrosylated thiol) to be converted back to the thiol to be identified. However, the assay here for H2S modification of the thiol has no such requirement. It is easy to undertake and has previously been used successfully to identify H2O2-sensitive proteins. Therefore, it is suggested here that with a modification of that assay, H2S-sensitive proteins may also be identified.

5. EXPERIMENTAL PROTOCOLS To identify H2S-sensitive proteins in cell samples, it is suggested that a modification of the methods used to previously identify H2O2-sensitive proteins is used (Hancock et al., 2005). However, previously this was used with plant samples, specifically from Arabidopsis thaliana. Here, to show that the method is applicable to a wider range of samples, Caenorhabditis elegans is used as a model system. Therefore, the methods below will outline the growth of the C. elegans, their treatment with H2S, the tagging of the thiol groups, and the separation of the subsequent proteins with polyacrylamide electrophoresis.

6. CAENORHABDITIS ELEGANS AS A MODEL ORGANISM Caenorhabditis elegans has been used as a model organism for many years, since Brenner suggested its use in 1965 (Brenner, 1974). It is a small nematode which is easy to grow, cheap to maintain, and has a short life span. But more importantly, the genome of C. elegans has been very well characterized, as have its developmental stages. Each worm is known to contain exactly 959 somatic cells, which to the large part have been characterized. Furthermore, because of the large amount of homology between genes in C. elegans and those in humans, these nematodes are very important in research on human disease and for drug discovery (Kaletta & Hengartner, 2006). For a full in-depth discussion on the use of C. elegans in laboratory work, refer to the WormBook (http://www.wormbook.org/). Interestingly, and of relevance here because the effects of H2S are being considered, it has been shown that treatment of C. elegans with H2S bestows on the worms a degree of thermal tolerance (Miller & Roth, 2007), and furthermore, the life span of the worms has been shown to be increased. More recently, it has been shown that C. elegans are capable of the generation of H2S, with the presence of three enzymes being reported (Qabazard et al., 2014), and alteration of the levels of H2S synthesizing enzymes altered

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the organism’s life span. H2S metabolism, therefore, appears to be part of the normal physiology of the worms. Clearly, therefore, there must be downstream responses to H2S in C. elegans, which may involve the modification of thiol groups on proteins. However, here C. elegans is used simply as a model system in which to show that the tagging and identification of proteins which may react with H2S can be achieved.

7. GROWTH OF C. ELEGANS C. elegans were grown on NGM plates (Nematode Growth Media plates) with E. coli OP50 as a food source, or alternatively they were grown in liquid culture (again with E. coli as a source of food). However, here sufficient worms could be collected from NGM plates, dispensing with the need for the liquid cultures (see the WormBook (http://www. wormbook.org/)). – For NGM plates, 3 g NaCl, 17 g agar, 2.5 g peptone, and 975 mL H2O were mixed, and subsequently autoclaved and cooled. To this, the following additions were made: 1 mL 1 M CaCl2, 1 mL 5 mg/mL cholesterol in ethanol, 1 mL 1 M MgSO4, 25 mL 1 M KPO4 buffer [108.3 g KH2PO4, 35.6 g K2HPO4, H2O to 1 L]. Large square petri dishes (120  120  17 mm) are ideal for growing worms. Plates were poured, and when ready were spread with E. coli using a glass rod. After incubation at 37 °C for at least 6 h or overnight at room temperature, the plates were cooled to room temperature and the worms added. Basically, worms are added by “chunking,” that is, chunks of NGM plate containing worms and E. coli are removed from one plate and added to a new plate, allowing the worms to multiple on a new food source. Alternatively, worms and E. coli were washed from a well-populated plate, briefly centrifuged at 1500  g, and then after removal of some of the supernatant the remaining sample can be pipetted onto a fresh E. coli plate. E. coli ideally should be grown at 37 °C but a 20 °C incubator should be used for growing the worms.

8. TREATMENT OF SAMPLES WITH H2S H2S is a gas and so samples can be treated directly with H2S from the gaseous phase (gas tanks of H2S can be purchased, e.g., from Praxair). If whole organisms are to be treated, or parts of an organism, such as the leaves

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of the plant, this may be a viable option, although strict safety measures will be needed due to the toxicity of H2S. However, the use of gas is not always convenient and suitable for treating biological samples, such as cells and cell cultures, or protein samples. Therefore, more often than not compounds which release H2S are used—so-called donor molecules. Common donors include sodium hydrosulfide (NaHS) and sodium sulfide (Na2S). These are cheap, easy to obtain and store, and easy to use. An example of such as study is one looking at apoptosis in animal cells, mediated by mitogen-activated protein kinases (Adhikari & Bhatia, 2008). But the caution that needs to be exercised here is that once in solution both these compounds will release H2S quickly, and therefore, samples will be exposed to relatively high H2S for a short period of time. To overcome this, new donor compounds have been designed and made available such as GYY4137 (Li et al., 2008), which release compounds in a more physiological manner, that is, more slowly for longer periods. Other donors have been generated that are targeted to organelles and are commercially available (from VIVA Biosciences). Very recently, a new H2S-releasing compound that is designed to be targeted to the mitochondria, AP-39, has been reported, with the view that it may lead to a new therapeutic strategy for a range of diseases (Le Trionnaire et al., 2014). A further discussion of such compounds can be found in this issue of Methods in Enzymology. However, here NaHS was used as a demonstration of the effects which can be seen. If other, and more recent, donors are used, it will be necessary to change the concentrations and time periods of treatment used.

9. ESTIMATION OF TOXICITY OF H2S COMPOUNDS H2S is extremely toxic, so care needs to be taken to ensure the effects of treatment with H2S donors are what they appear to be. For example, if investigating signaling aspects of H2S physiology, then it is important to be sure that there is a physiological response to H2S but that the organisms or cells are not simply being killed. Even though low concentrations of H2S can be used by cells as a source of reducing power for mitochondria (Bouillaud et al., 2013), H2S is also a potent inhibitor of mitochondrial function (Dorman et al., 2002) and so treatments need to be of concentrations which do not have this latter effect. Therefore, when using new H2S donors, or new organisms or tissues, investigations into the effects of H2S need to be carried out before isolation and analysis of protein samples. Here, the

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toxicity and physiological responses to the H2S donor NaHS are used as an example. – To assess the toxicity of a H2S donor on the nematodes, the worms were washed from NGM plates with M9 buffer (3 g KH2PO4, 6 g Na2HPO4, 5 g NaCl, 1 mL 1 M MgSO4 in 1 L of water, sterilized by autoclaving). – A specified amount of this liquid culture was used in each treatment, giving a final volume of 2 mL for each experiment. NaHS was added at the following concentrations: 0 M (M9 buffer control), 10 μM, 50 μM. Three repeats were carried out for each concentration of NaHS used. Nematodes were observed using a light microscope, and numbers showing slowed, normal, and no movement (presumed dead: that is, they appeared to be both not moving and stiff ) were counted using click counters at 10-min intervals up to 30 min. It could be argued that assessing movement of the worms is quite subjective, but it becomes obvious with experience. Furthermore, slowed movement is often accompanied by jerky movements, making it easier to assess. – The data for NaHS-treated nematodes are shown in Fig. 3. Here, the percentage of the worms showing reduced movement is shown. There was no significant toxicity during the period of this experiment (data not shown), but with other organisms, or H2S donors, toxicity may well be seen with this range of concentrations and the H2S donor may well be needed to be significantly diluted before use. – With several H2S donors, there will also be by-products formed on the release of H2S. Therefore, toxicity experiments, as well as protein analysis, should be repeated with time-depleted donors. That is, donor compounds should be incubated in the absence of biological material for the same periods of time as the experiment and then subsequently used. Such depletion experiments are routinely used when NO donors are employed for example, and the practice should be repeated here.

10. TREATMENT OF SAMPLES WITH THIOL TAG Samples which can be treated can be either whole organisms, whole tissues, cell cultures, or protein extracts. Clearly, if proteins are to be tagged and identified in whole organisms or tissues, issues around the penetration of the tag have to be considered. It may be preferable to label proteins while they are in their physiological environment, but this may not always be possible. Therefore, often proteins are isolated and treated and labeled in vitro, allowing the possible modifications of thiol groups on proteins to be

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Percentage of nematodes showing slowed movement

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Figure 3 Changes in C. elegans movement after treatment with fast-release H2S donor NaHS. Nematodes were washed from NGM plates with M9 buffer. A specified amount of this liquid culture was used in each treatment, giving a total volume of 2 mL for each repeat. Three repeats were carried out for each concentration of NaHS as well as an M9 buffer control. After addition of NaHS nematodes were observed using a light microscope, and numbers showing slowed, normal, and no movement (presumed dead) were counted using click counters at 10 min intervals up to 30 min. ddd 0 μM (buffer control); d•d•d 10 μM NaHS; • • • • 50 μM NaHS. Error bars show the 95% confidence interval around the mean for each sample. *Significant differences between samples (p < 0.05 by t-test).

estimated. Subsequently, such work would need to be repeated with the treatment, i.e., H2S, with the proteins in situ. This is the approach used here. Basically, worms were treated, protein extracted, and then tagged with 50 IAF for identification (see the scheme in Fig. 2). – To label protein samples, 50 -iodoacetamide fluorescein (IAF) was used. A stock solution needs to be made fresh before each use (5.15 mg in 500 μL PBS, pH 7.5). Stock solutions should be kept in the dark (wrapped in foil) before use. – Labeling the proteins is carried out by the addition of up to 200 μM IAF (final concentration) for 10 min before analysis. Again, samples need to be kept in the dark before analysis (covered in foil or placed in a dark place). Longer incubation times may be required if using whole cells or tissues. Here, to 200 μL of sample, 2 μL of 20 mM IAF was added and incubated at room temperature in the dark, before analysis by electrophoresis.

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11. ISOLATION AND ANALYSIS OF MODIFIED PROTEINS The final treated protein samples need to be analyzed by SDSpolyacrylamide gel electrophoresis. Samples are either treated with IAF or treated with H2S donors and then IAF needs to be denatured and separated. – For analysis of proteins capable of binding IAF in the example given here, proteins were isolated from the nematodes before treatment. However, it would be possible to treat whole worms or tissues with IAF before analysis. – For protein isolation, 10 large square petri dishes (120  120  17 mm) of worms were washed with 5 mL S basal buffer per plate (S basal buffer consists of 5.85 g NaCl, 1 g K2HPO4, 6 g K2PO4, 1 mL cholesterol (5 mg/mL in ethanol) in a liter of H2O). This was dispensed into 2  50-mL Falcon tubes, and the worms were pelleted at 3000 rpm for 2 min. The supernatant was discarded. Worms were washed in buffer and repelleted as before. – Samples were treated as required, with concentrations of H2S donors dictated by the results of the toxicity assay. Here 0–50 μM NaHS was used as a demonstration. Samples were treated for 20 min. – Worms were removed from the H2S donor by centrifugation (max 4.4  g) for 5 min, and the supernatant discarded. Worms were washed in S basal buffer and were subsequently resuspended in enough iced triethylammonium bicarbonate buffer (TEAB buffer, 100 mM (diluted from 1 M stock, pH 8.5)) to bring the volume up to approx. 0.4 mL. This was agitated for 20 s to aid lysis (use a “tissue ruptor” if available), followed by 30 s on ice. This was repeated three times and then centrifuged at 13.3  g for 2 min to remove unlysed material. – 200 μL supernatant was removed to new 1.5-mL Eppendorf tubes (50 μL was saved for BCA assay: this can be frozen for later analysis if needed for convenience). – 50 -IAF treatment was carried out as this point. Discussion and details can be found above. – The proteins were precipitated with acetone for further analysis. 1 mL ice cold acetone was added to the sample, which was vortexed and then incubated at 20 °C for 10 min. This was then pelleted at 13.3  g for 5 min and the acetone discarded. The sample was air dried for approximately 10 min. The sample was still kept covered as much as possible to reduce the amount of light it is exposed to.

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kDa

1

2

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2

Figure 4 5-IAF labeling of control and treated C. elegans protein samples. Protein was extracted from nematode samples either untreated or treated with 50 μM NaHS for 20 min. Protein samples were labeled with 5-IAF and separated using SDS-PAGE. Marker sizes are shown. Lane 1: Precision Plus Protein Standards (Bioline, Hertfordshire, UK); Lane 2: Control—protein from untreated nematodes [2.39 mg/mL]; Lane 3: Protein from nematodes treated with 50 μM NaHS [2 mg/mL]. A representative example of a typical gel is shown.

– To prepare the sample for electrophoresis, the pellet was resuspended in 20 μL sample loading buffer. 2 μL 10 DTT was added and mixed. The sample was denatured at 70 °C for 10 min and then separated on an acrylamide gel. The electrophoresis was subsequently run at 200 V, 100–125 mA for 30 min to 1 h (checking the progress of the dye front). – IAF-tagged proteins were then be visualized under UV light. The gel needs to be removed from between the plates and can then be viewed and photographed. An example can be seen in Fig. 4. – To visualize all the proteins on the gel, and not just those that have bound to the IAF, the gel needs to be stained for the presence of all proteins using Coomassie blue (approximately 30 min), with a methanol-based destain (30% for approximately 6 h). The gel can then once again be photographed, this time using visible light. This will indicate both the presence of non-IAF labeled proteins and allow comparison of the amount of protein present in different lanes on the same gel or across gels.

12. ESTIMATION OF PROTEIN CONCENTRATIONS IN SAMPLES Once samples have been isolated and are ready for treatment, it will be important to carry out a protein assay, as samples will need to be compared, either as different lanes on the same gel or across different gels.

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– Protein concentrations were estimated using a Pierce BCA Assay Protein Assay Kit (Thermo Scientific). This is a detergent-compatible assay which is based on bicinchoninic acid (BCA). It has long been known that there is a reduction of Cu2+ to Cu+ by protein in an alkaline medium (the so-called Biuret reaction). BCA can then be used to create a sensitive colorimetric detection of the cuprous cations (Cu+). Two molecules of BCA react with one Cu+ resulting in a purple reaction product. This can then be measured at 492 nm, either with a spectrophotometer or a plate reader. – Bovine serum albumin (BSA) should be used to create a standard curve against which unknown samples can be compared. BSA should be used as a serial dilution from 0.1 to 1 mg/mL. BSA was dissolved in TEAB buffer so that the buffer is consistent with the samples being assayed.

13. FURTHER ANALYSIS AND IDENTIFICATION OF MODIFIED PROTEINS Here, the labeling of protein samples from C. elegans was used as an example, and they were separated by 1-dimensional SDS-PAGE. It is clear from Fig. 4 that many proteins were labeled with IAF, showing that in a crude protein extract from the nematodes there are numerous proteins which have thiol groups accessible under the conditions used in this experiment. However, on pretreatment with H2S then many of these proteins are no longer visible under UV light, suggesting that they have bound to H2S rendering them incapable of subsequently binding to the IAF. However, this does not lead to the identification of these proteins. There are a variety approaches which may subsequently be used. – To determine which proteins have bound IAF, if the identification of the protein is either already known, or suspected, then an antibody approach may be used. Here the proteins can either be identified using Western blot analysis or an immunoprecipitation approach. – Alternately, an antibody against fluorescein can be used. This was the rationale used by Wu, Kwon, and Rhee (1998), for example. Here they treated samples with 5-IAF and then probed for the presence of the fluorescein tag with an antifluorescein antibody. One of the H2O2-sensitive proteins they identified was protein tyrosine phosphatase 1B, verifying the approach that they used. – Regardless of the alternatively method chosen the best way to identify the proteins is by separating them by 2-dimensional electrophoresis. The first dimension will be a separation due to the proteins’ isoelectric points, and

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different gradient and ranges can be used for this. If a complete set of proteins is to be analyzed a wide pH range will need to be employed but often to get better separate of proteins that appear close together a narrow pH range is used. The second dimension will be separation by molecular weight, and again the exact separation seen will depend on the percentage of acrylamide used in the gel. A standard of 10–12% is often used, but acrylamide gels may be more (up to about 15%) or less (down to approximately 5%) or indeed a gradient gel may be used, where the acrylamide gets progressively more concentrated down the length of the gel. – Once the proteins have been separated by 2-D electrophoresis, then the proteins need to be identified. This is often achieved using mass spectrometry. Protein spots are removed, often manually with a scalpel, and then the proteins are cleaved into peptides. This is usually done with trypsin. Peptides can be identified using matrix-assisted laser desorption ionization time-of-flight spectrophotometry. This yields a peptide mass fingerprint which can be compared to theoretical peptides derived from nucleotide databases, as the cut sites of trypsin can be determined. Alternately, if this does not yield a match, then MS/MS can be used to break the peptides and do further analysis for their identification. – Even once identified it cannot be assumed that the protein had reacted with H2S. Ideally further analysis needs to be carried out. This may include the identification of a mass shift in a peptide containing a cysteine residue, to show the exact product of the H2S reacting with the protein. Further enzyme analysis may also include the assay of enzyme activity (or protein function) in the presence and absence of varying concentrations of H2S or H2S donors.

14. CONCLUDING REMARKS The ability to measure the modifications of thiols is very important to get a full understanding of how reactive compounds such as H2S interact with proteins to alter their activities and functions. However, other compounds such as ROS, NO, and glutathione also have the ability to reactive with thiols to give altered proteins and peptides. Therefore, it has to be considered that H2S is not working on its own but rather in concert with a host of other reactive chemicals (Hancock & Whiteman, 2014). Therefore, similar investigations of thiol modifications needs to be carried out with other compounds, perhaps with other assays too, such as the biotin switch assay (Forrester et al., 2009; Haldar & Stamler, 2013; Nakamura et al., 2013;

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Zhang et al., 2005), as it is under the physiological conditions in which a protein resides that will determine the exact end result of the thiol alteration. If NO is the predominant signal, then perhaps S-nitrosylation will be the result, but if H2O2 is predominant then the thiol may be oxidized—to varying degrees. Getting a full picture of the possible thiol modifications that may take place, considering some thiols may not be solvent accessible, or be used for disulfide formation, and how such thiols are modified in a cell is therefore important. Such discussion also highlights that doing these assays in vitro is not always very representative of the true picture, so such experiments of treatments with NO donors, H2S donors, H2O2, etc. need to be carried out on whole cells or whole tissues/organisms too. Lastly, these experiments need to be carried out with combinations of reactive compounds too, so that any competition between such compounds is unraveled. It has been suggested that H2S, or H2S donor molecules, may be used as a future therapeutic agent (Xu et al., 2014) with specific examples including during inflammation (Fox et al., 2012). Furthermore, with plants the storage of fruit has been shown to be affected by the presence of H2S (Hu et al., 2012), the mechanism being suggested as alteration of antioxidant concentrations within the fruits. Therefore, the interest in the biology of H2S is surely going to increase, and the direct reacts that H2S has with biomolecules is going to be important to determine in a range of organisms, from bacteria and plants to animals, and in human tissues.

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